Formal study of CLS includes a curriculum of diverse natural and physical science courses in addition to the corps of minimum graduation requirements for a B.A. or B.S. with minor in Biology or Chemistry. Click here for a typical undergraduate curriculum for a BS degree in CLS.
In addition, registration with a professional association or licensure by many states requires a minimum 1 year internship in an accredited Medical Technology program provided at many larger hospital laboratories. Several years ago hospitals would pay a stipend to students in their programs. Most such programs now charge a tuition comparable to standard university tuition.
There are many basic procedures and methods which are common to nearly all clinical laboratories. Listed below are some common procedures (and some not so common) and their descriptions. Additional specialty methods (particularly immunological) are described here. A Glossary of Diagnostic Terminology is located here.
Brightfield Microscopy
With a conventional bright field microscope, light from an incandescent source is aimed toward a lens beneath the stage called the condenser, through the specimen, through an objective lens, and to the eye through a second magnifying lens, the ocular or eyepiece.
Specimens that may be observed using bright-field microscopy:
Prepared slides, stained - bacteria (1000x), thick tissue sections (100x, 400x), thin sections with condensed chromosomes or specially stained organelles (1000x), large parasites (100x). Smears, stained - blood (400x, 1000x), stained bacteria (400x, 1000x). Living preparations (wet mounts, unstained) - yeast suspensions (40x, 100x, 400x), living parasites or their ova (40x, 100x, 400x ccasionally), fungal preperations (40x, 100x, 400x).
It is best to start with the lowest magnification objective lens, to home in on the specimen and/or the part of the specimen you wish to examine. It is rather easy to find and focus on sections of tissues, especially if they are fixed and stained, as with most prepared slides. However it can be very difficult to locate living, minute specimens such as bacteria or unpigmented cells.
The lowest power lens is usually 3.5 or 4x, and is used primarily for initially finding specimens. We sometimes call it the scanning lens for that reason. The most frequently used objective lens is the 10x lens, which gives a final magnification of 100x with a 10x ocular lens. For very small details in prepared slides such as cell organelles, you will need a higher magnification. Typical high magnification lenses are 40x and 97x or 100x. The latter two magnifications are used exclusively with oil in order to improve resolution.
Move up in magnification by steps. Each time you go to a higher power objective, re-focus and re-center the specimen. Higher magnification lenses must be physically closer to the specimen itself, which poses the risk of jamming the objective into the specimen. Be very cautious when focusing. Good quality sets of lenses are parfocal, that is, when you switch magnifications the specimen remains in focus or close to focused.
Adjust illumination so that the field is bright without hurting the eyes. Adjust the condenser. The condenser lense is used to focus light on the specimen through an opening in the stage. Start with the aperture diaphragm stopped down (high contrast). You should see the light that comes up through the specimen change brightness as you move the aperture diaphragm lever.
Dark Field Microscopy
To view a specimen in dark field, an opaque disc is placed underneath the condenser lens, so that only light that is scattered by objects on the slide can reach the eye. Instead of coming up through the specimen, the light is reflected by particles on the slide. Everything is visible regardless of color, usually bright white against a dark background. Pigmented objects are often seen in "false colors," that is, the reflected light is of a color different from the color of the object. Better resolution can be obtained using dark as opposed to bright field viewing.
You don't need sophisticated equipment to get a dark field effect, but you do need a higher intensity light, since you are seeing only reflected light. At low magnification (up to 100x) any decent optical instrument can be set up so that light is reflected toward the viewer rather than passing through the object directly toward the viewer.
| Phase Contrast Microscopy Most of the detail of living cells is undetectable in bright field microscopy because there is too little contrast between structures with similar transparency and no color. Unless the specimen mount is extremely thin, dark field mode may distort details. However, the various organelles show wide variation in refractive index, that is, the tendency of the materials to bend light, providing an opportunity to distinguish them. Highly refractive structures bend light to a much greater angle than do structures of low refractive index. The same properties that cause the light to bend also delay the passage of light by a quarter of a wavelength or so. In a light microscope in bright field mode, light from highly refractive structures bends farther away from the center of the lens than light from less refractive structures and arrives about a quarter of a wavelength out of phase. Light from most objects passes through the center of the lens as well as to the periphery. If the light from an object to the edges of the objective lens is retarded a half wavelength and the light to the center is not retarded at all, then the light rays are out of phase by a half wavelength. They cancel each other when the objective lens brings the image into focus. A reduction in brightness of the object is observed. The degree of reduction in brightness depends on the refractive index of the object. Phase Contrast Condensers and Objective Lenses To use phase contrast the light path must be aligned. An element in the condenser is aligned with an element in a specialized phase contrast lens. This usually involves sliding a component into the light path or rotating a condenser turret. The elements are either lined up in a fixed position or are adjusted by the observer until the phase effect is optimized. Generally, more light is needed for phase contrast than for corresponding bright field viewing, since the technique is based on a diminishment of brightness of most objects. |  Schematic configuration for phase contrast microscopy: Light passing through the phase ring is first concentrated onto the specimen by the condenser. Undeviated light enters the objective and is advanced by the phase plate before interference at the rear focal plane of the objective. Photomicrograph of hair cross sections from a fetal mouse using phase contrast microscopy (200X). |
| Polarized Light Microscopy Many transparent solids are optically isotropic, meaning that the index of refraction is equal in all directions throughout the crystalline lattice. Examples of isotropic solids are glass, sodium chloride, many polymers, and a wide variety of both organic and inorganic compounds. Crystals are classified as being either isotropic or anisotropic depending upon their optical behavior and whether or not their crystallographic axes are equivalent. All isotropic crystals have equivalent axes that interact with light in a similar manner, regardless of the crystal orientation with respect to incident light waves. Light entering an isotropic crystal is refracted at a constant angle and passes through the crystal at a single velocity without being polarized by interaction with the electronic components of the crystalline lattice. Anisotropic crystals have crystallographically distinct axes and interact with light in a manner that is dependent upon the orientation of the crystalline lattice with respect to incident light. When light enters non-equivalent axes, it is refracted into two rays, each polarized with their vibration directions oriented at right angles to each other and traveling at different velocities. One of the rays travels with the same velocity in every direction through the crystal and is termed the 'ordinary ray'. The other ray travels with a velicity that is dependent upon the propagation direction within the crystal and is termed the 'extraordinary ray'. A polarizer is placed beneath the substage condenser causing polarized light to enter the anisotropic crystal where it is refracted into two separate component rays vibrating parallel to the crystallographic axes and perpendicular to each other. The polarized light waves then pass through the specimen and objective lense before reaching a second polarizer, termed the 'analyzer', that is oriented to pass a polarized vibration direction perpendicular to that of the substage polarizer. |  Schematic configuration for polarized light microscopy: White light is plane polarized by the substage polarizer and concentrated onto the anisotropic specimen by the condenser. Light rays emerging from the specimen interfere when they are recombined in the analyzer, subtracting some of the wavelengths of white light and producing a myriad of colors. Photomicrograph of high-density liquid crystalline calf thymus DNA using polarized light microscopy (100X). |
Microscopy with Oil Immersion
When light passes from a material of one refractive index to material of another, as from glass to air or from air to glass, the light bends. Light of different wavelengths bends at different angles, so that as objects are magnified the images become less and less distinct. This loss of resolution becomes very apparent at magnifications of above 400x or so. Even at 400x the images of very small objects are badly distorted.
Placing a drop of oil with the same refractive index as glass between the cover slip and objective lens eliminates two refractive surfaces and considerably enhances resolution, so that magnifications of 1000x or greater can be achieved.
To use an oil immersion lens, first focus on the area of specimen to be observed with the high dry (400x) lens. Place a drop of immersion oil on the cover slip over that area, and very carefully swing the oil immersion lens into place. Focus carefully, preferably by observing the lens itself while bringing it as close to the cover slip as possible, then focusing by moving the lens away from the specimen. When in focus, the lens nearly touches the cover slip. The focal plane is so narrow that it is very easy to focus right past it. If you are focusing toward the specimen, you can drive the lens right into it.
When to Use Oil Immersion Objective Lenses
Use an oil immersion lens when you have a fixed (dead - not moving) specimen that is no thicker than a few micrometers. Even then, use it only when the structures you wish to view are quite small - one or two micrometers in dimension. Oil immersion is essential for viewing individual bacteria or details of the striations of skeletal muscle. It is nearly impossible to view living, motile microorganisms at a magnification of 1000x, except for the very smallest and slowest.
A disadvantage of oil immersion viewing is that the oil must stay in contact, and oil is viscous. A wet mount must be very secure to use oil. Oil immersion lenses are used only with oil, and oil can't be used with dry lenses, such as your 400x lens. Lenses of high magnification must be brought very close to the specimen to focus and the focal plane is very shallow, so focusing can be difficult. Oil distorts images seen with dry lenses, so once you place oil on a slide it must be cleaned off thoroughly before using the high dry lens again. Oil on non-oil lenses will distort viewing and possibly damage the coatings.
Fluorescence Microscopy
To understand how fluorescence microscopy works and why it has become so important to modern biology, one must understand what the term fluorescence means. Fluorescence is the luminescence of a substance when it is excited by radiation. In microscopy, fluorescence is used as a means of preparing specific biological probes. Some biological substances like chlorophyll and some oils and waxes have primary fluorescence; that is, they autofluoresce. But most biological molecules or structures do not fluorescence on their own, so they must be linked with fluorescent molecules (or fluorochromes) in order to create specific fluorescent probes.
| Fluorescence of a substance is seen when the molecule is exposed to a specific wavelength of light (excitation wavelength or spectrum) and the light it emits (the emission wavelength or spectrum) is always of a higher wavelength. To view this fluorescence in the microscope, several light filtering components are needed. Specific filters are needed to isolate the |  |
| One other component is required: a dichroic beam splitter or partial mirror which reflects lower wavelengths of light and allows higher wavelengths to pass. A beam splitter is required because the objective acts as a condenser lens for the excitation wavelength as well as the objective lens for emission. One only wishes to see the light emitted from the fluorochrome and not any of the excitation light, and the beam splitter isolates the emitted light from the excitation |  |
UV and Visable Spectrophotometry
Spectrophotometry is the use of an instrument called a spectrophotometer to detrmine absorption spectra of compounds, perform kinetic assays or to determine the concentration of organic or inorganic analytes in solution. The instrument may produce tunable or fixed monochromatic light of specific wavelength for absorbance measurements. The most common adaptation uses wavelengths of the visible light portion (ca. 340-900 nanometers - 1 nm=10-9 Meter) of the electromagnetic spectrum. For operation between 340-1000 nm, a tungsten filament lamp is sufficient; for operation between 180-340 nm the light source must be a Hydrogen or Deuterium lamp and the cuvette cells must be quartz glass.
 Diagram of the basic components of a spectrophotometer, consisting of light source (Lamp), Prism or grating, Colimator, Cuvette (Test Cell), Detector (selenium or silicon photocell or photomultiplier tube) and Display (analog or digital). In some implementations the Lamp, Prism and Colimator are a separate unit known as a 'Monochromator'. Physical arangements and enhancements vary with specific applications. |  Spectronic 20 Spectrophotometer: It is manufactured in several configurations. |
The Beer-Lambert Law
Within limitations, the law states that when a sample is placed in the beam of a spectrophotometer, there is a direct linear relationship between the amount (concentration) of its constituent(s) and the amount of energy it absorbs. This may be stated mathematically:
or rearranged,
where,
I0 = incident radiation, I = transmitted radiation
A = absorbance
e = extinction coefficient at a given wavelength
l = pathlength in cm.
C = molar concentration, [M]
(The extinction coefficient, molar absorptivity, has units M-1cm-1; hence, most standard cuvette cells have a pathlength of 1 cm.)When the molar absorptivity constant is known for a substance at a specific wavelength, the Beer equation may be used to directly determine concentration. Example: NADH (reduced nicotinamide adenine dinucleotide), a substance employed in many enzyme-catalized chemical reactions, is known to have a molar absorptivity of 6.220 mM-1cm-1 at 340 nm. Measurement of the absorption of an unknown concentration of NADH at 340 nm and application of the Beer equation will yield a quantitative measurment of NADH in solution. For example, if the absorption of an unknown NADH solution at 340 nm in a 1 cm cell is 0.622, then the concentration may be calculated from
Cu = 0.622/[(6.22 mM-1cm-1)(1 cm)] = 0.1 mM NADH
Of course, the utility of this measurement is predicated on the knowledge that this is a pure solution of NADH or that there are no other 340 nm-absorbing substances present.
More commonly, however, an analyte must be measured by indirect photometric methods. This circumstance arises when the molar absorptivity of a substance is unknown or when the absorbance of a substance is very low at all practical spectral wavelengths or when there are other substances present which absorb at the substance's spectral maxima.
An indirect photometric method employs a chemical reaction(s) to produce another substance which has unique absorption properties. A procedure which uses this method is the simplified Fearon analysis for urea, a waste product of protein catabolism which is produced in the liver and excreted by the kidney. While urea in solution is
 In the Fearon procedure for serum urea quantitation, urea condenses with diacetyl to form diazine. Since diacetyl is unstable, diacetyl monoxime is substituted and generates the required diacetyl in the same reaction mixture. Also, thiosemicarbazide and ferric ions are added to enhance and stabalize the product. | virtually transparent to all practical visible and uv wavelengths, the new product, diazine, absorbs strongly at 540 nm and its concentration is directly related to the concentration of urea. Calibration Calibration of a method is applied using a wide variety of mathematical methods or mathematical models. In practice, nearly all photometric methods require calibration with standard solutions of known value because of |
where Cu = Concentration of the Unknown solution
Cs = Concentration of the Standard solution
Au = Absorbance of the Unknown solution
As = Absorbance of the Standard solution
INFRARED SPECTROSCOPYIR is one of numerous spectrometric techniques for analyzing the chemistry of materials. In all cases, spectrometric analysis implies a measurement of a very specific wavelength of light energy, either in terms of amount absorbed by the sample in question, or the amount emitted from the sample when suitably energized.
| IR is an absorption form of spectrometric analysis. Unlike atomic absorption, IR is not concerned with specific elements (such as Lead, Copper, etc.) but, rather, with the groupings of atoms in specific combinations to form what are called "functional groups". These various functional groups help to determine a material's properties or expected behavior. By knowing which wavelengths are absorbed by each functional group of interest one can cause the appropriate wavelength to be directed at the sample |  |
FOURIER TRANSFORM INFRARED SPECTROSCOPY
Fourier transform infrared (FTIR) spectroscopy is a powerful analytical tool for characterizing and identifying organic molecules. The IR spectrum of an organic compound serves as its fingerprint and provides specific information about chemical bonding and molecular structure.
Microbeam FTIR allows areas as small as 10-15 microns to be analyzed; this allows the source of organic particles to be determined. Using attenuated total reflectance (ATR), thin films can be analyzed directly on a surface. SSL uses FTIR to aid in precise determination of the chemical identity of organic contamination in a variety of samples including samples from the disk drive, biomedical, semiconductor, PCB, electronic, laser and optic industries.
ADVANTAGES OF FTIR:
Small spot size (10-15 microns)
Detailed chemical bonding information
Organic analysis of polymers and plastics
Analysis of liquids, solids, and gases
In ATR mode can sample the outer ~ 1000
Non-destructive analysis
Can analyze non-conductive materials
Molecular specific identification
RAMAN SPECTROSCOPY
Raman spectroscopy is based on the inelastic scattering of light by a molecule, and is a type of spectroscopy that is complementary to infrared spectroscopy. There are two common methods for measuring vibrational spectra. Infrared spectroscopy is a direct method which looks at the transition between vibrational levels in a single electronic state. IR transitions range from < 200 cm-1 to >4000 cm-1.
IR spectroscopy can be a useful tool for identifying molecules; however, its usefulness is limited by several factors:
1) A molecule must have a change in its dipole moment upon absorption if the absorption is to take place. Thus, molecules such as homonuclear diatomics are IR inactive. 2) In general, transitions between a vibrational level and the next higher vibrational level (v -> v+1) are strong.
Some of these problems are reduced in Raman spectroscopy. This type of spectroscopy is based on light scattering. If light of a particular wavelength is aimed at a sample, much of the light will pass through, but some will be scattered in all directions. Of that scattered light, most comes out at the same wavelength it went in: no energy is exchanged between the light and the molecule. This is elastic or Rayleigh scattering. But some light will lose energy to the molecule; however, it can only lose an amount of energy that is the same as one of the vibrational transitions in the molecule. Therefore, the scattered light has less energy (a longer wavelength) than the incoming light. This is inelastic or Raman scattering. In order to see the Raman scattering, one must detect away from where the incident light shines through. The detector includes a monochromator which allows us to determine the spectrum of the scattered light.
Raman and IR spectroscopy are complementary and overlapping techniques for understanding molecular rotations and vibrations. A large IR absorption requires a large change in the electric dipole moment of the molecule [mij =
yi x yj dv >> 0]. A large Raman signal requires a large change in the polarizability of the molecule [aij = yi ayj dv >>0 ]. While many transitions have both large mij and large aij, there are many that don't. Therefore, Raman and IR can act together to give a more complete description of the molecule.Chromatography is a separation process involving two phases, one stationary and the other mobile. Typically, the stationary phase is a porous solid (e.g., glass, silica, or alumina) that is packed into a glass or metal tube or that constitutes the walls of an open-tube capillary. The mobile phase flows through the packed bed or column. The sample to be separated is injected at the beginning of the column and is transported through the system by the mobile phase. In their travel through the column, the different substances distribute themselves according to their relative affinity for the two phases. The rate of travel is dependent on the values of the distribution coefficients, the components interacting more strongly with the stationary phase requiring longer time periods for elution (complete removal from the column). Thus, separation is based on differences in distribution behaviour reflected in different migration times through the column. At the present time, chromatography is the most significant method for separation of organic substances and, along with electrophoresis, is most widely used for biological substances.
The various chromatographic methods are characterized in terms of the mobile phase--gas: gas chromatography (GC); liquid: liquid chromatography (LC); supercritical fluid: supercritical-fluid chromatography (SFC). The methods are then further subdivided in terms of the stationary phase; thus, if the stationary phase is a solid adsorbent, there are methods such as gas-solid chromatography (GSC) and liquid-solid chromatography (LSC). Chromatography is conducted with computer-controlled instrumentation for high precision and unattended operation. In addition, a detector is frequently placed on-line after the column for either structure analysis or quantitation or both. One of the most powerful approaches of analysis now available is the on-line coupling of chromatography to mass spectrometry.
 In Gas Chromatography, the determining factor in how fast a component travels is usually (but not always) the boiling point of the compound. (If a polar high-boiling liquid adsorbent is used in the GC column, the polarity of the components determines the elution order.) | Gas chromatography is an important method owing to its speed, resolving power, and detector sensitivity. Since it depends on vaporization, this technique is best suited to compounds that can be vaporized without suffering decomposition. Many substances that normally do not easily vaporize can be chemically derivatized for successful volatilization separation by gas chromatography. Right-click and click Reload to view the animation at left. |
Since the early 1970s, liquid chromatography has developed as the premier separation method for organic substances. Because the mobile phase is a liquid, the requirement for vaporization is eliminated, and therefore LC can separate a much broader range of substances than GC. Species that have been successfully resolved include inorganic ions, amino acids, drugs, sugars, oligonucleotides, and proteins. Both analytical-scale liquid chromatography with samples at the microgram-to-milligram level and preparative-scale liquid chromatography at the tens-of-grams level have been developed. In biotechnology, preparative-scale liquid chromatography is especially important for purification of proteins and peptide hormones made by recombinant technology.
One important method is liquid-solid chromatography in which the porous adsorbent is polar, and separation is based on the properties of classes of compounds--e.g., amines (alkaline) from alcohols (neutral) and esters (neutral) from acids. Liquid-solid chromatography is the oldest of the chromatographic methods. Until the mid-20th century, the experimental procedure had not changed much from its original form. After significant improvements, liquid-solid chromatography now is conducted with porous particles as small as 3-5 micrometres (0.00012-0.00020 inch) in diameter, and liquid pumps are used to drive the liquid through the particle-filled column. High resolution and fast separations are achieved since the small particles allow good efficiency with fast mobile phase velocities (one centimetre per second or higher). This technique is also important in purification, and separated substances can be automatically collected after the column using a fraction collector.
A significant liquid-solid chromatography procedure is reverse-phase chromatography, in which the liquid mobile phase is water combined with an organic solvent such as methanol or acetonitrile and the stationary phase surface is nonpolar or hydrocarbon-like. In contrast to normal-phase chromatography, where the adsorbent surface is polar, in reverse-phase chromatography the elution of substances from the column is in the order of increasing polarity. In addition, separation is based on the nonpolar aspects of the substances. In the separation of a series of peptides from human growth hormone, a recombinantly made drug, an enzyme, trypsin, is used to break peptide bonds containing the basic amino acids--arganine and lysine--to yield a specific fingerprint of the protein. Peptide mapping is a critical method for evaluating the purity of complex substances such as proteins.
Ion-exchange chromatography (IEC) is a subdivision of liquid-solid chromatography, but its importance is such that it deserves special mention. As the name implies, the process separates ions; the basis of the separation is the varying attraction of different ions in a solution to oppositely charged sites on a finely divided, insoluble substance (the ion exchanger, usually a synthetic resin). In a cation-exchange resin all the sites are negatively charged, so that only positive ions can be separated; an anion-exchange resin has positively charged sites. Ion-exchange chromatography has become one of the most important methods for separating proteins and small oligonucleotides. An important application of ion exchange is the removal of dissolved iron, calcium, and magnesium ions from hard water. The negative sites on a cation exchanger are first neutralized with sodium ions by exposure to a strong solution of common salt (sodium chloride); when the hard water is passed through the resin, the undesirable ions in the water are replaced by sodium ions.
Liquid-solid adsorption chromatography also can be performed on thin, flat plates (thin-layer chromatography, or TLC). TLC is inexpensive and rapid but not as sensitive or efficient as column chromatography. In practice, the adsorbent is spread on a glass plate and dried. The sample is applied as a spot near one end of the plate, which is placed (vertically) in a shallow reservoir containing the mobile phase. As the mobile phase travels up the plate by capillary action, the sample dissolves in the liquid, and its components are transported up the plate to new positions at varying distances from the starting point.
Separation of chemical components of a mixture is achieved due to the selective interaction of chemicals with both the stationary and mobile phases:
In Gas Chromatography, the determining factor in how fast a component travels is usually (but not always) the boiling point of the compound. (If a polar high-boiling liquid adsorbent is used in the GC column, the polarity of the components determines the elution order.)
In Column and Thin Layer chromatographies, the stationary phase (the adsorbent: silica gel or alumina) is polar, and the polarities of both the component of the mixture and the solvent used as the mobile phase are the determining factors in how fast the compound travels.
Column chromatography is used to separate and purify components of a mixture. TLC and GC are usually (but not always!) used only to analyze mixtures: to determine the number of components and to see if a desired component is present. TLC is often used to determine the "ideal system" for a column chromatography procedure.
Determining solvent systems for TLC and Column Chromatography
When you need to determine the best system (a "system" means the eluting solvent, itself often a mixture of solvents) to develop a TLC plate or chromatography column loaded with an unknown mixture, vary the polarity of the solvent in several trial runs -- a process of trial and error. Carefully observe and record the results of the chromatography in each solvent system. You will find that as you increase the polarity of the solvent system, all the components of the mixture move faster (and visa versa with lowering the polarity). The ideal solvent system is simply: the system that separates the components.
TLC elution patterns usually extrapolate to column chromatography elution patterns. Since TLC is a much faster procedure than column chromatography, TLC is often used to determine the best solvent system for column chromatography. For instance, in determining the solvent system for a flash chromatography procedure, the ideal system is the one that moves the desired component of the mixture to a TLC Rf of 0.25-0.35 and will separate this component from its nearest neighbor by difference in TLC Rf values of at least 0.20. Therefore a mixture is analyzed by TLC to determine the ideal solvent(s) for a flash chromatography procedure.
Beginners often do not know where to start: What solvents should they pull off the shelf to use to elute a TLC plate? Because of toxicity, cost, and flammability concerns, the common solvents are hexanes (or petroleum ethers, ligroin) and ethyl acetate (an ester). Diethyl ether can be used, but it is very flammable and volatile. Alcohols (methanol, ethanol) can be used. Acetic acid (a carboxylic acid) can be used, usually as a small percentage component of the system, since it is corrosive, non-volatile, very polar, and has irritating vapors. Acetone (a ketone) can be used. Methylene chloride (halogenated hydrocarbon) is a good solvent, but it is toxic and should be avoided whenever possible. If two solvents are equal in performance and toxicity, the more volatile solvent is preferred in column chromatography because it will be easier to remove from the desired compound after isolation from a column chromatography procedure. Mix a non-polar solvent (hexanes, a mixture of 6-carbon alkanes) with a polar solvent (ethyl acetate or acetone) in varying percent combinations to make solvent systems of greater and lesser polarity.
Coupling Gas Chromatography to Mass Spectrometry
The suite of gas chromatographic detectors includes (roughly in order from most common to the least): the flame ionization detector (FID), thermal conductivity detector (TCD or hot wire detector), electron capture detector (ECD), photoionization detector (PID), flame photometric detector (FPD), thermionic detector, and a few more unusual or VERY expensive choices like the atomic emission detector (AED) and the ozone- or fluorine-induced chemiluminescence detectors. All of these except the AED produce an electrical signal that varies with the amount of analyte exiting the chromatographic column. The AED does that AND yields the emission spectrum of selected elements in the analytes as well. Another GC detector that is also very expensive but very powerful is a scaled down version of the mass spectrometer. When coupled to a GC the detection system itself is often referred to as the mass selective detector or more simply the mass detector. This powerful analytical technique belongs to the class of hybrid analytical instrumentation (since each part had a different beginning and can exist independently) and is called gas chromatograhy/mass spectrometry (GC/MS).
Placed at the end of a chromatographic column in a manner similar to the other GC detectors, the mass detector is more complicated than, for instance, the FID because of the mass spectrometer's complex requirements for the process of creation, separation, and detection of gas phase ions. A capillary column is most often used in the chromatograph because the entire MS process must be carried out at very low pressures (~10-5 torr), and in order to meet this requirement, a vacuum is maintained via constant pumping using a vacuum pump. It is difficult for packed GC columns to be interfaced to an MS detector because they have carrier gas flow rates that cannot be as successfully pumped away by normal vacuum pumps; however, capillary columns' carrier flow is 25 or 30 times less and therefore easier to "pump down." That said, GC/MS interfaces have been developed for packed column systems that allow for analyte molecules to be dynamically extracted from the carrier gas stream at the end of a packed column and thereby selectively sucked into the MS for analysis. For one type interface, using a silicone membrane, the selectivity for organic molecules (the analyte) over helium (the carrier gas) is 50,000.
The high cost for the pump, ionization source, mass filter or separator, ion detector, and computer instrumentation and software has limited the wide application of this system as compared to the less expensive GC detectors (e.g., FID cost ~$3000 vs. MS cost ~$40,000). However, the power of this technique lies in the production of mass spectra from each of the analytes detected instead of merely an electronic signal that varies with the amount of analyte. These data can be used to determine the identity as well as the quantity of unknown chromatographic components with an assuredness simply unavailable by other techniques.
Components of the GC/MS
| Leaving the entire capillary GC system aside, the major components of the mass selective detector itself are: an ionization source, mass separator, and ion detector. There are two common mass analyzers or separators commercially available for GC/MS; they are the quadrapole and the ion trap. Right-click and click Reload to view the animation at right. In this example, the lightest fragment is B+; the heaviest A+. The last frame of the movie is a mass spectrum displaying only these three fragments. Their relative mass to charge ratios are specified by their relative position on the x axis (low mass/charge to left, high mass/charge to right). The relative amounts (commonly called peak intensity) of each of these fragments determined during the mass analyzer's scan is reflected on the y axis. GC/MS is a reference method used in the clinical laboratory to detect, identify and quantitate DAU (Drugs of AbUse) and by forensic toxicology laboratories. Click here for example IR and Mass spectra (Cocaine). |  The GIF animation is a short series of steps for the process of a single analyte (already separated from the other analytes in the chromatographic mixture) denoted as ABC exiting the chromatographic column and: the analyte (A-B-C) undergoing ionization and fragmentation; the charged fragments (A+ B+ C+) being separated by mass; the fragments which are focused on the mass filter's exit slit passing into the detector; and the charged ions being detected. |
Flow cytometry is a method for quantitating components or structural features of cells primarily by optical means. Although it makes measurements on one cell at a time, it can process thousands of cells in a few seconds. Since different cell types can be distinquished by quantitating structural features, flow cytometry can be used to count cells of different types in a mixture.
 | Flow cytometers have been commercially available since the early 1970's, and their use has been increasing since then. The most numerous flow cytometers are those used for complete blood cell counts in clinical laboratories -- these do not employ fluorescence. More versatile research instruments employ fluorescence, hence may be distinguished as flow cytofluorometers. < Screen print of Coulter MAXM hemogram. Cell types are classified by size, volume and internal structure. |
Flow cytofluorometers are found in all major biological research institutions. They are also numerous in medical centers, where they are used for diagnosis as well as research. There are about 7,000 flow cytofluorometers in use worldwide. Ploidy and cell cycle analysis of cancers is the major diagnostic use. Lymphomas and leukemias are intensively studied for surface markers of diagnostic and prognostic value. Although less expensive alternative technologies are under development, until the present time, flow cytometry has been the method of choice for monitoring CD4 lymphocyte levels in the blood of AIDS patients.
The term "FACS" is Becton-Dickinson's registered trademark and is an acronym for Fluorescence-Activated Cell Sorter. Another major vendor of flow cytometers is Coulter Electronics.
The cells may be alive or fixed at the time of measurement, but must be in monodisperse (single cell) suspension. They are passed single-file through a laser beam by continuous flow of a fine stream of the suspension. Each cell scatters some of the laser light, and also emits fluorescent light excited by the laser. The cytometer typically measures several parameters simultaneously for each cell:
low angle forward scatter intensity, approximately proportional to cell diameter
orthogonal (90 degree) scatter intensity, approximately proportional to the quantity
of granular structures within the cell
fluorescence intensities at several wavelengths
Light scatter alone is often quite useful. It is commonly used to exclude dead cells, cell aggregates, and cell debris from the fluorescence data. It is sufficient to distinguish lymphocytes from monocytes from granulocytes in blood leukocyte samples.Fluorescence intensities are typically measured at several different wavelengths simultaneously for each cell. Fluorescent probes are used to report the quantities of specific components of the cells. Fluorescent antibodies are often used to report the densities of specific surface receptors, and thus to distinguish subpopulations of differentiated cell types, including cells expressing a transgene. By making them fluorescent, the binding of viruses or hormones to surface receptors can be measured. Intracellular components can also be reported by fluorescent probes, including total DNA/cell (allowing cell cycle analysis), newly synthesized DNA, specific nucleotide sequences in DNA or mRNA, filamentous actin, and any structure for which an antibody is available. Flow cytometry can also monitor rapid changes in intracellular free calcium, membrane potential, pH, or free fatty acids.
Flow cytometers involve sophisticated fluidics, laser optics, electronic detectors, analog to digital converters, and computers. The optics deliver laser light focused to a beam a few cell diameters across. The fluidics hydrodynamically focus the cell stream to and within an uncertainty of a small fraction of a cell diameter, and, in sorters, break the stream into uniform-sized droplets to separate individual cells. The electronics quantitate the faint flashes of scattered and fluorescent light, and, under computer control, electrically charge droplets containing cells of interest so that they can be deflected into a separate test tube or culture wells. The computer records data for thousands of cells per sample, and displays the data graphically.
Analysis Equipment: the FACScan
The Becton-Dickinson FACScan is used for analysis of cell samples. Unlike the sorter (see below), this instrument cannot separate cells into different containers based on their properties; the samples are consumed and discarded during analysis. The FACScan is a closed fluidic system, so use with biohazardous samples (such as human blood samples) is possible with appropriate precautions.
The FACScan is easy to use, and the instrument is operated by the experimenter her/himself. The FACScan uses an air-cooled argon gas laser, 15 mW output, with a fixed wavelength emission of 488 nm. It has three fluorescence detection channels which simultaneously detect green, yellow-orange, and red light. Fluorescein is used extensively for the green channel, and phycoerythrin or propidium iodide (a DNA stain) for the yellow-orange channel. Dyes are also available which can be excited at 488 nm yet emit in the red.
The FACScan can analyze cell suspensions at the rate of several hundred cells per second. Typically, investigators acquire 5,000 to 15,000 cells per sample. Data are saved to the hard disk of a dedicated Hewlett-Packard computer, where they can later be analyzed with graphics software. Data can also be transferred to a network server computer so that they are accessible from any computer network. Excellent public domain software is available for PC's which may be copied freely to any computer for data analysis.
Sorting Equipment: the FACStar Plus
A Becton-Dickinson FACStar Plus is used for cell sorting, and for analysis requirements which cannot be met on the FACScan. This instrument has a class IV water-cooled argon gas laser with a rated output in all-wavelength mode of 4 watts. This laser can be tuned, so the FACStar is used, for example, when the analysis requires excitation at 514 nm in addition to 488 nm.
The FACStar Plus can sort cells, or acquire data, at a rate of several thousand per second (about 10-fold faster than the FACScan analyzer). Because the FACStar uses a stream-in-air sorting method, it aerosolizes the sample and cannot be used for biohazardous samples. Nonhazardous living cells can be sorted, and may be recovered in gnotobiotic ("sterile") form for subsequent in vitro functional studies.
Enzyme Structure
Enzymes are extraordinarily efficient and selective biological catalysts that accelerate the chemical reaction toward equilibrium. Chemical reactions in the metabolic pathways necessary for the maintenance of life would not proceed at reasonable rates without enzymes. Enzymatic reactions are 103to 1017 times faster than the corresponding uncatalyzed reactions.
Enzymes are globular protein molecules (M.Wt. typically 104 - 105 ) that catalyze the myriad of biochemical reactions that occur within living cells. Like their chemical counterparts, enzymes accelerate the rate of chemical reactions without themselves being changed in the overall process. A fundamental difference between enzymes and industrial catalysts however is that enzymes function at physiological temperatures in a low ionic strength solution at near neutral pH. There are many different kinds on enzyme, each promoting only a very limited range of chemical reactions. Enzymes are very efficient catalysts. One molecule of catalase for example can decompose 40,000 molecules of hydrogen peroxide per second at freezing point.
The table below gives turnover numbers for some other enzymes at room temperature.
Enzyme Turnover number
(molecules/second)
Carbonic anhydrase 600,000
Acetylcholinesterase 25,000
amylase 18,000
Penicillinase 2,000
DNA polymerase 15
The substrate molecules are bound in a hydrophobic cleft know as the active site. Enzymes whose activity is regulated generally have a more complex structure than unregulated enzymes. Most regulatory enzymes have two sites - one for the substrate, and the other for the modulator.Enzyme Kinetics
An important feature of enzymes is that they possess specific 3-D configurations that are fundamental to their biological function. This is because the overall shape of the molecule stabilizes the precise geometric structure of the active site, the region in the enzyme where the substrate is converted into the product. The importance of the active site (which normally makes up only a small percentage of the entire molecule) is to stabilize the transition state between the substrate and its products thus lowering the activation energy for the reaction. (Lowering this energy by about 34 kJ mol-1 is calculated as bringing a million fold increase in the rate of a reaction at 298 K.) For this to occur the substrate must fit precisely into the active site (shape recognition). Indeed, the "lock and key" analogy is often used to describe substrate binding, as substrate fits an enzyme like a key fits a lock. The activation energy for the enzyme catalyzed reaction can be determined in the normal way by the Arrhenius equation;
Each enzyme requires certain definite conditions for optimum performance, particularly as regards to pH, temperature and ionic strength. The presence of specific accessory substances (co-factors, activators etc.) may also be a requirement.
Enzymes are unstable substances, and are easily inactivated by high temperature or extremes of pH. Although they are not consumed during the biochemical reaction, enzymes are inherently labile and have to be continuously synthesized.
For many enzyme reactions the rate (V or dp/dt) varies with the substrate concentration [S] as shown below. The rate (V or dp/dt) is defined as the number of moles per second which is a measure of enzyme activity. At low substrate concentration the rate or velocity (V) is almost proportional to substrate[S] concentration. At high substrate [S] concentrations, the velocity is not linear with the [S] concentration and rate approaches a maximum velocity called Vmax.
The following model was proposed by Michaelis and Menten in 1913 to explain the kinetics of an enzyme reaction (equation 1). One of first great advances in biochemistry was the discovery that an enzyme transiently binds to a substrate to form a enzyme-substrate complex [ES]. The substrate binds noncovalently to the active site of the enzyme. The rate of an enzymatic reaction depends on the concentrations of both the substrate and the catalyst (enzyme). When the amount of enzyme is much less than the amount of substrate, the reaction is pseudo first order. The straight line illustrates the effect of enzyme concentration on reaction velocity in pseudo first-order reaction. The more enzyme present, the faster the reaction.
 | Pseudo first-order conditions are used in analyses that determine the concentration of an enzyme in a sample. The concentration of enzyme can be easily determined by comparing its activity to a reference curve as shown at left.Modern automated clinical chemistry analyzers store the kinetic reaction constants in their computer and calculate unknown values from measured absorbance changes over standard time. This implementation of enzyme measurement is known as a kinetic method. In fact, most analyses (not merely for enzymes) employ a kinetic method - because they are faster. End-point methods often require up to an hour or more for the reaction to achieve equilibrium, whereas kinetic methods may require just a few minutes. |
At the beginning of an enzyme-catalyzed reaction, the amount of product formed is negligible, and the reaction can be described by equation 1. Note that the conversion of the ES complex into free enzyme and product is shown by a one-way arrow. During the initial period when measurements are made little product has been formed, so the rate of the reverse reaction is small. The velocity measured during this short period is called the initial velocity (vo). The use of initial velocity simplifies the interpretation of kinetic data and avoids complications that may arise as the reaction progresses, such as product inhibition, depletion of substrate, and slow denaturation of the enzyme.
Equation 2:
Equation 3:
Equation 5:
Equation 6:
Equation 7:
The meaning of Km is evident from the Michaelis-Menten equation. When [S] = Km, then V=Vmax/2. Thus, Km is equal to the substrate concentration at which the reaction rate is half of its maximal value.
The Michaelis constant, Km, and the maximal rate, Vmax, can be readily derived from rates of catalysis measured at different substrate concentrations if an enzyme operates according to the simple scheme given in the Michaelis Menten equation. It is convenient to transform the Michaelis-Menten equation into one that gives a straight line plot. This can be done by taking the reciprocal of both sides of the equation.
A plot of 1/V versus 1/[S], called a Lineweaver-Burk plot, yields a straight line with and intercept of 1/Vmaxand slope of Km/Vmax
Figure 2.
Figure 1: The plot is the reaction velocity V verses the concentration of the substrate [S]. The maximum velocity Vmax is the velocity where the enzyme is saturated with substrate[S] and 1/2 Vmax is that substrate [S] where the velocity 1/2 Vmax.
The Michaelis constant, Km, and the maximal rate, Vmax can be readily derived from rates of enzyme catalyzed reaction measured at different substrate concentrations. The enzyme must operate according to the simple scheme given in equation 1.
It is convenient to transform the Michaelis-Menten equation into one that gives a straight line plot. This can be done by taking the reciprocal of both sides of the equation. A double-reciprocal plot of enzyme kinetics: 1/V is plotted as a function of 1/[S]. The slope is Km/Vmax, the intercept on the vertical axis is 1/Vmax, and the intercept on the horizontal axis is -1/Km.
Km and Vmax for an enzyme-catalyzed reaction can be measured in several ways. Both values can be obtained by analysis of initial velocities at a series of substrate concentrations and a fixed concentration for the enzyme. In order to obtain reliable values for the kinetic constants, the [S] point must be spread out both below and above Km to produce a hyperbola. It is difficult to determine either Km or Vmax directly from a graph of initial velocity versus concentration because the curve approaches Vmax asymptotically. However, using a suitable computer program, accurate values can be determined by fitting the experimental results to the equation for the hyperbola.
The Michaelis-Menten equation can be rewritten in order to obtain values for Vmax and Km from straight lines on graphs. The most commonly used transformation is the double-reciprocal Lineweaver-Burk plot. Values of Km can be determined even when enzymes have not been purified, provided that only one enzyme in the impure preparation can catalyze the observed reaction.
Table 2 Km values of some enzyme
| Enzyme | Substrate | Km (mM) |
| Chymotrypsin | Acetyl-L-tryptophanamide | 5,000 |
| Lysozyme | Hexa-N-acetylglucosamine | 6 |
| b-Galactosidase | Lactose | 4,000 |
| Threonine deaminase | Threonine | 5,000 |
| Carbonic anhydrase | CO2 | 800 |
| Penicillianase | Benzylpenicillin | 50 |
| Pyruvate Carboxylase | Pyruvate HCO3- ATP | 400 1,000 50 |
| Arginine-tRNA synthetase | Arginine tRNA ATP | 3 0.4 300 |
At high substrate concentration, when [S] is much greater than Km, V=Vmax, i.e., the rate is maximal, independent of substrate concentration. When V=Vmax then [S]/([S] + Km) approaches 1. When [S] = Km then V = Vmax/2. Thus Km is equal to the substrate concentration at which the reaction rate is half of its maximal value. When the [S] is less than Km then the V=Vmax [S]/Km and the substrate concentration is proportional to the velocity.
The significance of Km and Vmax values
The K m values of enzymes range widely (0.4 µM for Arginine-tRNA synthetase to 5,000 µM for chymotrypsin). For most enzymes Km lies between 10-1 and 10-7 M. The Km value for am enzyme depends on the particular substrate and also on environmental conditions such as pH, temperature, and ionic strength. The Michaelis constant, Km has two meanings. First, Km is the concentration of substrate at which half the active sites are filled. Once the Km is known, the fraction of site filled at any substrate concentration can be calculated from the equation
Second K m is related to the rate constants of the individual steps in the catalytic scheme given in the following equation.
Km=(k2 + kcat)/k1
Equation 1:
K m =k2 /k1
The dissociation constant of the ES complex is given by
K ES =[E][S]/[ES]=k2 /k1
In other words, Km is equal to the dissociation constant of the ES complex if kcat is much smaller than k2. When this condition is met. Km is a measure of the strength of the ES complex: A high Km indicates weak binding ; a low Km indicates strong binding. It must be stressed that Kmindicates the affinity of the ES complex only when k2 is much greater than kcat.
Turnover number
The turnover number of an enzyme is the number of substrate molecules converted into product by an enzyme molecule in a unit of time when the enzyme is fully saturated with substrate. It is equal to the kinetic constant kcat. The maximal rate Vmax, reveals the turnover number of an enzyme if the concentration of active site [ET] is known, because
V max =kcat [ET]
For example, a 10-6 M solution of carbonic anhydrase catalyzes the formation of 0.6 M H2CO3 per second when it is fully saturated with substrate. Hence, kcat is 6 X 105 s-1.
kcat= Vmax/Et = 0.6/10-6=6 X 105 S-1
This turnover number is one of the largest known. Each round of catalysis occurs in a time equal to 1/kcat, which is 1.7 microseconds for carbonic anhydrase. The turnover numbers of most enzymes with their physiological substrates fall in the range from 10 to 107 per second.
| Enzyme | Kcat(s-1) |
| Papain | 10 |
| Ribonuclease | 100 |
| Carboxypeptidase | 100 |
| Trypsin | 100 to 1,000 |
| Acetylcholinesterase | 1,000 |
| Kinases | 1,000 |
| Dehydrogenases | 1,000 |
| Transaminases | 1,000 |
| Carbonic anhydrase | 1,000,000 |
| Superoixe dismutase | 1,000,000 |
| catalase | 10,000,000 |
Kinetic Perfection in Enzymatic Catalysis (The K cat /K m)
When the substrate concentration is much greater than Km, the rate of catalysis is equal to k cat, the turnover number, as described in the preceding section. However, most enzymes are not normally saturated with substrate. Under physiological conditions, the [S]/Km ratio is typically between 0.01 and 1.0. when [S]<<Km, the enzymatic rate is much less than kcat because most of the active sites are unoccupied. Is there a number that characterizes the kinetic of an enzyme under these conditions? Indeed there is, as can be shown by combining two equations.
This restriction also pertains to enzymes having more complex reaction pathways. Their maximal catalytic rate when substrate is saturating, denoted by kcat, depends on several rate constants rather than on kcat alone. The pertinent parameter for these enzymes is kcat/Km. In fact, the kcat/Km ratios of the enzymes acetycholinesterase, carbonic anhydrase, and trisephophate isomerase are between 108 and 109 M-1 S-1, which shows that they have attained kinetic perfection. Their catalytic velocity is restricted only by the rate at which they encounter substrate in the solution. Any further gain in catalytic rate can come only by decreasing the time for diffusion. Indeed, some series of enzymes are associated into organized assemblies so that the product of one enzyme is very rapidly found by the next enzyme. In effect, products are channeled from one enzyme to the next, much as in an assembly line.
The constants kcat and kcat.Km are useful for comparing the activities of different enzymes. It is also possible to assess the efficiency of an enzyme by measuring the rate acceleration that it provides. This value is the ratio of the rate constant for a reaction in the presence of the enzyme (kcat) divided by the rate constant for the same reaction in the absence of enzyme(kn). Surprisingly few rate acceleration values are known because most cellular reactions occur extremely slowly in the absence of enzymes.
Competitive and Noncompetitive Inhibition
Measurements of the rates of catalysis at different concentrations of substrate and inhibitor serve to distinguish between competitive, uncompetitive and noncompetitive inhibition. An inhibitor (I) is a compound that binds to an enzyme and interferes with its activity by preventing either the formation of the ES complex or its breakdown to E +P. Inhibitors are used experimentally to investigate enzyme mechanisms and to decipher metabolic pathways. Natural inhibitors regulate metabolism, and many drugs are enzyme inhibitors. Inhibition can be either irreversible or reversible. Irreversible inhibitors are bound to enzymes by covalent bonds. Reversible inhibitors are bound to enzymes by the same noncovalent forces that bind substrates and products.
The constant for the dissociation of I from the EI complex, called the inhibition constant (Ki), is described by the equation Ki =[E][I]/[EI]
Effects of Reversible Inhibitors on Kinetic Constants
| Type of inhibitor | Effect |
| Competitive (I bind to E only) | Raises Km and Vmax remains unchanged |
| Uncompetitive (I binds to ES only | Lowers Vmax and Km Ratio of Vmax.KmRemains unchanged |
| Noncompetitive (I binds to E or ES) | Lowers Vmax Km remains unchanged |
Competitive Inhibition
When a nonmetabolizable molecule, I, resembles a metabolizable molecule, S, sufficiently to be bound to the enzyme, then I remains attached to the enzyme and prevents the attachment of S, thus competing with S for space on the enzyme surface. The inhibition of the reactions of S is called classical competitive inhibition. Nonclassical competitive inhibition is the binding of S at the active site and prevents the binding of I at a different site. The reverse is also true.
Ethanol is Used Therapeutically as a Competitive Inhibitor to Treat Ethylene Glycol Poisoning.
About fifty deaths occur annually from the ingestion of ethylene glycol, a constituent of antifreeze. Ethylene glycol itself is not lethally toxic. Rather, the harm is done by oxalic acid, the oxidation product of ethylene glycol, because the kidneys are severely damaged by the deposition of oxalate crystals. The first committed step in this conversion is the oxidation of ethylene glycol to an aldehyde by alcohol dehydrogenase. This reaction can be effectively inhibited by administering a nearly intoxicating dose of ethanol. The basis of this effect is that ethanol is a competing substrate and so it blocks the oxidation of ethylene glycol to aldehyde products. The ethylene glycol is then excreted harmlessly. Ethanol is also used as a competing substrate for treating methanol poisoning.
In competitive inhibition, the intercept of the plot of 1/V versus 1/[S] is the same in the presence and absence of inhibitor, although the slope is different. This reflects the fact that Vmax is not altered by a competitive inhibitor. Competitive inhibition can be overcome by a sufficiently high concentration of substrate. At a sufficiently high concentration, virtually all the active sites are filled by substrate and the enzyme is fully operative. The increase in the slope of the 1/v versus 1/[S] plot indicates the strength of binding of competitive inhibitor. In the presence of a competitive inhibitor the Lineweaver-Burk equation becomes:
In other words, the slope of the plot is increased by the factor (1+[I]/ki) in the presence of a competitive inhibitor. Consider an enzyme with a Km of 10-4M. In the absence of inhibitor, V=Vmax/2 when [S]=10-4 M. In the presence of 2 X 10-3 M competitive inhibitor that is bound to the enzyme with a Ki of 10-3 M, the apparent Km will be 3 X 10-4M. Substitution of these values into Lineweaver Burk equation give V=Vmax/4.
Noncompetitive Inhibition
In some cases the inhibitor resembles the metabolizable substance, but it is apparently attached to the enzyme at some point other than the one which binds the original substrate, and by its antagonistic action it prevents the expected enzyme reaction.
In noncompetitive inhibition Vmax is decreased to Vimax, and so the intercept on the vertical axis is increased. The new slope, which is equal to Km/VImax, is larger by the same factor. In contrast with Vmax, Km is not affected by this kind of inhibition. Noncompetitive inhibition cannot be overcome by increasing the substrate concentration. The maximal velocity in the presence of a noncompetitive inhibitor is Vimax.
Noncompetitive inhibitors probably alters the conformation of the enzyme to a shape that can still bind S but cannot catalyze any reaction by forming product.
.
Uncompetitive inhibitors bind only to ES, not to free enzyme. In uncompetitive inhibition, Vmax is decreased by the conversion of some molecules of E to the inactive form ESI. Since it is the ED complex that binds I, the decrease in Vit1 is not reversed by the addition of more substrate. Uncompetitive inhibitors also decrease the Km because the equilibria for the formation of both ES and ESI are shifted toward the complexes by the binding of I. Experimentally, the lines on a double-reciprocal plot representing varying concentrations of an uncompetitive inhibitor all have the same slope, indicating proportionally decreased value for Km and Vmax. This type of inhibition usually occurs only with multisubstrate reactions.
There are approximately 3000 enzymes which have been characterized. These are grouped into six main classes according to the type of reaction catalyzed. At present, only a limited number are used for analytical purposes.
Oxidoreductases
These enzymes catalyze oxidation and reduction reactions involving the transfer of hydrogen atoms or electrons. The following are of particular importance in the design of enzyme electrodes. This group can be further divided into 4 main classes.
Dehydrogenases catalyze hydrogen transfer from the substrate to a nicotinamide
adenine dinucleotide cofactor (NAD+). An example of this is lactate
dehydrogenase which catalyzes the following reaction:
Lactate + NAD+ = Pyruvate + NADH + H+
Oxidases catalyze hydrogen transfer from the substrate to molecular oxygen
producing hydrogen peroxide as a by-product. An example of this is FAD
dependent glucose oxidase which catalyses the following reaction:
b-D-glucose + O2 = gluconolactone + H2O2
Peroxidases catalyze oxidation of a substrate by hydrogen peroxide. An
example of this type of enzyme is horseradish peroxidase which catalyzes the
oxidation of a number of different reducing substances (dyes, amines,
hydroquinones etc.) and the concomitant reduction of hydrogen peroxide. The
reaction below illustrates the oxidation of neutral ferrocene to ferricinium in the
presence of hydrogen peroxide:
2[Fe(Cp)2] + H2O2 + 2H+= 2[Fe(Cp)2]+ + 2 H2O
Oxygenases catalyze substrate oxidation by molecular oxygen. The reduced
product of the reaction in this case is water and not hydrogen peroxide. An
example of this is the oxidation of lactate to acetate catalyzed by
lactate-2-monooxygenase.
lactate + O2 = acetate + CO2 + H2O
TransferasesThese enzymes transfer C, N, P or S containing groups (alkyl, acyl, aldehyde, amino, phosphate or glucosyl) from one substrate to another. Transaminases, transketolases, transaldolases and transmethylases belong to this group.
Hydrolases
These enzymes catalyze cleavage reactions or the reverse fragment condensations. According to the type of bond cleaved, a distinction is made between peptidases, esterases, lipases, glycosidases, phosphatases and so on. Examples of this class of enzyme include; cholesterol esterase, alkaline phosphatase and glucoamylase.
Lyases
These enzymes non-hydrolytically remove groups from their substrates with the concomitant formation of double bonds or alternatively add new groups across double bonds.
Isomerases
These enzymes catalyse intramolecular rearrangements and are subdivided into;
racemases
epimerases
mutases
cis-trans-isomerases
An example of this class of enzyme is glucose isomerase which catalyses the
isomerisation of glucose to fructose.
LigasesLigases split C-C, C-O, C-N, C-S and C-halogen bonds without hydrolysis or oxidation. The reaction is usually accompanied by the consumption of a high energy compound such as ATP and other nucleoside triphosphates. An example of this type of enzyme is pyruvate carboxylase which catalyzes the following reaction:
Prosthetic groups have similar function to co-factors with the exception that they are tightly bound to the enzyme. When they are released, the enzyme is mostly denatured. Flavin nucleotides and hemes are the most important examples of this class of molecule.
Listed below are examples of co-factors and prosthetic groups.
Compound Function
Nicotinamide adenine dinucleotide (NAD+) hydrogen transfer
Flavin mono nucleotide hydrogen transfer
Flavin adenine dinucleotide (FAD) hydrogen transfer
Heme electron transfer
Ferredoxins electron transfer, hydrogen activation
The production of monoclonal antibodies (MAbs) was pioneered by Georges Kohler and Cesar Milstein at the laboratory of molecular biology at Cambridge. For this significant achievement they were awarded the Nobel Prize in Physiology or Medicine in 1984. The technique they developed allowed the production of large quantities of homogeneous antibody against almost any antigen. MAbs have been one of the most powerful and widely used technologies in both research and medicine. They were christened magic bullets.
What are Monoclonal Antibodies?
MAbs are all produced from descendents of a single ancestral hybridoma cell and are identical. They only recognize a single specific antigenic determinant or hapten. They are secreted continuously by hybridoma cells in culture. Antibodies are made up of 4 polypeptide chains (2 heavy and 2 light) and these make up the characteristic Y-shape. Below are 3 diagrams showing the very basic structure of an antibody and how it interacts with antigen.
Diagrams kindly reproduced from slides from Dr. Anne White
There are many advantages of MAbs which make MAbs useful in a number of applications.
They are of one defined specificity.
They can be produced indefinately by cells in culture.
Large quantities of antibody can be obtained from hybridomas grown in mice.
The immunogen need not be pure.
They cut down on the number of lab animals required.
The alternative to using MAbs is using polyclonal antisera, however this had a number of disadvantages: It is a mixture of antibodies of high and low affinity antibody populations.
It is a mixture of antibodies of different specificities.
The supply is limited to the life span of the immunized animal.
The quantity that is produced is small.
The immunogen has to be pure.
This limits the use of polyclonal antisera and advocates the use of MAbs.
Production of MAbsThe Original Method Devised By Kohler and Miltstein
Below is a diagram to summarize the method they developed. Click on different areas of the diagram to obtain further information.
Four basic steps were involved:
The generation of B-cell hybridomas by fusing antigen-primed B cells and myeloma cells.
Selection of the fused clones.
Screening of clones for antibody secretion with desired antigenic specificity.
Propagation of desired hybridomas.
Production of Human MAbsThere are many problems with using MAbs from different species like mice for applications in humans. The production of human antibodies has received much attention. Despite the efforts of many in this field, it has been difficult, but not impossible. Simply carrying out the same conventional method devised by Kohler and Milstein is not really an option. People are not able to be immunized with the same range of antigens as they have been able to do in the mouse or other species. To overcome this they have primed B cells in vitro, but the different microenvironment means that only low affinity IgM antibody is produced. One cannot just remove the spleen of a human, thus hybridomas must be prepared from peripheral blood, and this contains very few primed B cells. There is a lack of human myeloma cells that exhibit immortal growth. One must devise a different method to obtain human MAbs.
One way is to transform human B cells with the Epstein-Barr Virus (EBV) - which confers immortality to the cells. The lymphocytes are cultured with antigen in the presence of EBV. A small proportion will be immortal and secrete antibodies.
Another way is to incorporate the genes incoding human antibody in a SCID (Severe Combined Immunodeficiency) mouse. The mice can then be immunized and activated human B cells can be isolated from the spleen, but there is still the problem of a lack of human myeloma cells that can be used.
Finally, there is the option of producing engineered MAbs.
Production of Engineered MAbs
The traditional methods have not changed much since their original description. New technologies such as genetic engineering and computer modelling have made it possible to produce MAbs in completely novel ways. The undesirable consequences of using other species MAbs in humans could be eliminated by using engineered MAbs. Engineered MAbs could finally realize the full potential of these magic bullets.
The production of engineered MAbs can be divided into 2 main groups:
Chimeric and hybrid antibodies.
Phage display antibodies.
Chimeric and hybrid MAbsThese engineered antibodies are a combination of both human and mouse antibodies. They are commonly referred to as humanized. It allows MAbs to be produced of an identical specificity as well characterised mouse MAbs. Originally, the variable region sequences were cloned from a mouse antibody gene along with the genetic informaton needed for it to be expressed, plus the constant regions from human antibody gene. These are then combined and the engineered antibody is a human-mouse chimera. The antigenic specificity is determined by the mouse variable region DNA, while the isotype is determined by the human constant region DNA. It is now possible to produce chimeric anitbodies with just mouse CDRs via a process called 'CDR-grafting'. In this process the murine CDRs are grafted onto a Fc/Fv framework.
The engineered antibodies have fewer mouse antigenic determinants and are less likely to illicit an immune response. CDR-grafted antibodies have the least. This increases the chances for therapeutic uses of MAbs in humans. Another advantage is that the antibody retains the effector region from the human constant region.
Phage display
Phage display was developed at the beginning of the 1990's, and has revolutionized the generation of MAbs. The development of phage display has now made it possible to consider the isolation of human antibodies directly without immunization. Recent advances in the field of human immunogenetics and in phage technology have led to the assembly of "naive" human repertoires in vitro as complex as the natural immune system. In this method cDNA antibody variable regions are obtained and expressed as a part-molecule (FV) on the surface of a M13 filamentous phage. These phage are then used to infect bacteria, and the FV protein of defined specificty and affinity is secreted in large amounts into the culture medium.
Applications of MAbs
There is a large range of applications for monoclonal antibodies. These include:
Immunoassay and Immunodetection
Immunocytochemistry
Immunoprecipitation
Immunoblotting
Immunoaffinity Purification
ImmunoassayIn the process of immunoassay a factor is quantitated on the binding of specific antibody. The antibody is labelled with something that can be measured and then compared to a calibration curve. Widely used examples include ELISA (Enzyme-Linked-Immuno-Sorbent-Assay) and RIA (Radio-Immuno-Assay). For an assay to be successful it needs to be:
highly sensitive
have low non-specific binding
able to assay a large number of samples in a short time
ImmunocytochemistryImmunocytochemistry uses labelled antibody which binds specifically to a molecule of interest in a fixed cell or tissue mounted on a microscope slide. The antibody can be labelled with a number of things, commonly a fluorescent molecule, that can be used to visualize certain aspects of a cell like cell surface markers which cannot normally be distinguished. MAbs are used because their specificity is defined, and one may be sure when one uses MAbs that only the molecule of interest will be bound.
There are four steps involved:
Cell or tissue preparation
Fixation and preparation of thinly sliced sections on slides
Antibody binding
Detection
ImmunoprecipitationThis is the process in which antibody precipitates a factor which has been labelled and isolated from cells. It can also be used to identify antigens in a complex mixture, like immunoblotting. It is sometimes required because the antigen can become denatured and some of the epitopes destroyed during immunoblotting. Immunoprecipitation can be used with soluble antigens or with cell-surface antigens. The test antigen is labelled with I-125, the MAB is added and only binds to the one specific antigen and forms AB-Ag complexes. The complexes are precipitated out using a co-precipitating agent (e.g., Staphylococcal A) and spun out and washed to remove unbound antigens. The precipitate is resolubilized in a seperation gel and the protiens of the immune complex are separated. They are then autoradiographed to show the posistion of the bound antigen.
Immunoblotting
This process involves the characterization of a factor based on the MW and ability of antibody to bind to it. It is used to identify and characterize antigens from a complex mixture. The antigen samples are resolved by seperation in an analytical gel such as dodecyl sulphate (SDS), peptide mapping gels, or isoelectric focusing gels. The resolved molecules are transferred electrophoretically to a nitrocellulose membrane in a blotting tank. The blot is then treated with a MAb to the specific antigen in question. This is then washed and treated with a radiolabelled conjugate, which detects the bound antibodies. This is then washed again and put in contact with a X-ray film to produce an autoradiograph. The bands of antigen that have been bound by the MAbs are visible.
Immunoaffinity Purification
In this process an antibody on a column isolates a factor from a mixture. It allows the purification of a substance from 1,000 to 10,000-fold.
Image kindly reproduced from a slide from Dr. Anne White.
Uses of MAbs
The many uses for monoclonal antibodies can divided up into three main areas:
Research
Diagnostics
Therapeutics
ResearchMAbs have much potential for research in all areas of Biology. They allow one to detect and visualise cell components or molecules. For example, cell surface markers - such as those which helped to identify the different subsets of T cells. They can visualise the distribution of molecules in cells; for example they have been able to visualise the presence of the different types of molecule, e.g. actin, tubulin in the cytoskeletons of cells.
Diagnostics
MAbs have had most success in the field of diagnostics. MAbs can be used to detect even the very smallest quantities of a substance. Because of their high degree of specificity one can be sure that diagnostic tests that use MAbs will be highly accurate. The use of MAbs in diagnostics have allowed a high degree of standardization and precision. Examples of where they have been used as diagnostic tools:
Blood and tissue typing - important in blood transfusions and transplantation.
Quantification of T cell levels in HIV patients
A screening test for prostate cancer - The PSA test
Diagnosis of microbial infections - it identifies the organism involved
Pregnancy testing - MAbs detect the human chorionic gonadotrophin hormone
A screening test for HIV.
TherapeuticsMost excitement in this area has been the potential treatments for cancer, though so far there has been limited success. Immunotoxins are one of a new class of drugs undergoing clinical trials for the treatment of leukaemia. They are conjugated antibodies carrying a toxin such as ricin, and specifically target cancer cells. The toxin would inactivate the cell's ribosomes and inhibit protein synthesis. As the toxin does not attack the whole cell, only small amounts are required.
| One of the most common pieces of equipment used to separate materials into subfractions in a biochemistry lab is the centrifuge. A centrifuge is a device that spins liquid samples at high speeds and thus creates a strong centripetal force causing the denser materials to travel towards the bottom of the centrifuge tube more rapidly than they would under the force of normal gravity. Types of centrifuges. The major distinguishing features between centrifuge types are speed and capacity. In a typical biochemistry laboratory you will find three different centrifuges (this is true in your biochemistry teaching lab as well). The smallest are the so-called microfuge centrifuges. These are made for spinning 1 to 2 ml plastic centrifuge tubes at speeds up to 12 or 13 thousand rounds per minute. They have very small, light rotors in them (the rotor is the part of the centrifuge that contains the holes for the sample tubes) which speed up and slow down rapidly. These centrifuges are very convenient for low to medium speed centrifugation of small quantities of material. The next common size centrifuge is the large superspeed centrifuge. These have speeds up to about 20,000 rpm and can take tubes of various sizes, depending on the rotors (the larger the rotor, the slower the maximum speed). Typical tubes hold 25 or 30 mls but bottles as large as several hundred mls can be run with the correct rotor. Finally, most biochemistry laboratories have access to an ultracentrifuge. Speeds up to 70,000 rpm are available on typical modern versions. Again the size of tube and the maximum speed vary from rotor to rotor, but tube sizes up to about 60 mls are available. The theory behind centrifugation. The idea here is pretty straight forward and mechanical. If you want the more dense materials to be separated from the less dense materials, you need a force that differentiates between particles of different density. Think about a swimming pool with a rock and a piece of styrofoam. The rock is denser than water and thus it sinks. The styrofoam is less dense than water, and thus it floats. Density is of course mass per unit volume. So, if you have a bag full of rocks and styrofoam and you want to separate one from the other, just dump the mixture into some water under the influence of the earth's gravity. The rocks will displace the water because they have greater mass for a given volume and gravity will pull them through the water. On the other hand, the water will displace the styrofoam because a certain volume of water weighs more than the same volume of styrofoam. However, there are many things that are much closer in density than rocks and styrofoam and it is much harder to separate them just under the Earth's gravity. In addition, diffusion is always at work as random motion smears out small differences due to density. To overcome this, or sometimes just to make the separation process faster, it would be nice to come up with a way of generating larger mass (density) dependent forces than are available from the Earth's gravity alone. Another way to generate a mass dependent force is to spin something. As you know from physics, a body in motion tends to continue in motion along a straight path unless some force is exerted on it to change its path. Thus in order to force something to go in a circle, we must exert force on it pulling it in towards the center. An equal and opposite force will always result, pushing out from the center. This is cetripital force, and it is just the mass of the object times the acceleration required to keep it from flying outward along a straight line. Thus, things with larger mass (for a given volume) will have a greater force exerted on them and they will move towards the outer edge of the container more quickly than the things with a lower mass per volume. The acceleration required to keep the object from flying outward along a straight path is given by w2r where the greek letter omega stands for the speed of revolution (see below for units) and r is the distance from the axis of the revolution to the position of the sample. To get a the force felt by the sample, we multiply this by the mass of the sample: F = mw2r, where F is the force and m is the mass. Please notice two things about this equation. It is linearly proportional to the mass of the object, but the force increases with the square of the rotation speed. Thus, going from 1000 rpm to 10,000 rpm increases the forces involved by a factor of 100. Some practical considerations. So let's say I want to spin two 30 mls tubes of water at 50,000 rpm. What kinds of forces are involved? Well, for comparison, consider the force that the acceleration of gravity exerts on the 30 mls of water when it is sitting on a bench top. The acceleration of gravity is about 10 m/s2 (10 meters per second squared). The mass of 30 mls of water is about 30 g or about 0.03 kg. Thus the force involved at 1 g (1 times the acceleration of gravity) is 0.3 kg m/s2 or 0.3 Newtons. How about at 50,000 rpm? First we need to do some units conversion. 50,000 rpm is 50,000/60 = 830 rounds per second. Further, in order to make the units work out, we must convert rounds per second to radians per second (there are 2p radians in a complete round or circle so multiply by this factor). This gives 5200 radians per second (this is omega in the equation). The force is mw2r so the total force is 66,000 N (assuming that the sample is about 8 cm from the center of rotation). To get the number of times greater this is than gravity along, we divide by 0.3 (see above) and get about 220,000 g's. That means that the water sample spinning at 50,000 rpm is equivalent to a 13,000 lb truck at normal gravity. If you look inside an ultracentrifuge, you will find that the rotor (the thing that contains the samples) is sitting on a shaft. The shaft is rather small and actually wobbly. There is no way you could suspend a 13,000 lb truck from this shaft. So how does it work? It works because of symmetry. You always must have a sample directly across from the sample of interest that has the same overall mass. Thus, the two masses and the forces on them balance out and the shaft feels no torque. The moral of the story is that you must balance tubes for centrifuge runs. For ultracentrifuge runs, you must balance the tubes very, very carefully. I usually try to get the masses within about 0.03 grams for an ultracentrifuge run. Slower centrifuges you can use an accordingly less stringent balance procedure. Rotors. There are many types of rotors, but most fall into one of three classes: fixed angle rotors, swinging bucket rotors and vertical rotors. Most of the time you will used a fixed angle rotor. Occasionally a swinging bucket or vertical rotor with be used, but not very often any form (swinging bucket allows the sample to swing out to the horizontal, giving the longest possible path of movement of the particles. The vertical rotors do the opposite -- they are used when a very short overall path of centrifugation is required.Types of centrifugation applications. There are many ways in which centrifuges are used. More often than not they are used to sediment some material leaving the rest in solution. However, one can also use two other common applications for separating materials: equilibrium density sedimentation and kinetic density sedimentation. In the first case, the material is either layered on top of or mixed into some material that can either be preformed into a density gradient or will become a density gradient when it is spun at high speed. The centrifuge is then run until the material finds its place as a band of particular density within the tube. The kinetic density methods also generally involve long runs that allow the molecule to find a region of the medium with the same density and come to equilibrium. In kinetic density sedimentation, you do not run the gradient to the end. You start with a band of your sample on top of the tube and let it progress through the density gradient for some period of time. |
References
Enzyme Kinetics; Cardosi, Marco, Electrochemical Sensors Group; 2001; University of Paisley, Paisley, Scotland, U.K.
Enzyme Kinetics, Natural Toxins Research Center; 2001; Texas A&M University, Kingsville, TX
Flow Cytometry; Martz, E., Cote, C.; 2000; The University of Massachusetts, Amherst, MA
Optical Microscopy (Molecular Expressions Review), Davidson, M.W., Abramowitz, M.; 1999; Florida State University, Tallahassee, FL
The Beer-Lambert Law; The Squirer Group; 1999; The University of California, San Diego, CA
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